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Archived - Synopsis of Infectious Diseases and Parasites of Commercially Exploited Shellfish

Perkinsus marinus ("Dermo" Disease) of Oysters

Category | Common Name | Scientific Name | Distribution | Host Species
Impact on Host | Diagnostic Technique | Methods of Control | References | Citation


Category

Category 1 (Not Reported in Canada)

Common, generally accepted names of the organism or disease agent

Perkinsus marinus, "Dermo" Disease, Proliferative disease, Perkinsosis.

Scientific name or taxonomic affiliation

Perkinsus marinus (=Dermocystidium marinum, =Labyrinthomyxa marina). The genus Perkinsus was originally placed in the Order Perkinsida of the Class Perkinsea within the Phylum Apicomplexa (Levine 1978). However, the conoid structure in the zoospores (as described below) is incomplete suggesting that the apicomplexan affinity is tenuous (Villalba et al. 2004). Early taxonomic analysis based on nucleotide sequences indicate that Perkinsus is phylogenetically closer to dinoflagellates and to coccidean and piroplasm apicomplexans than to fungi or flagellates. More recent investigations indicate that this parasite may not belong in the Phylum Apicomplexa but seems to be more closely related to the Dinoflagellida (Goggin and Barker 1993, Perkins 1996, Siddall et al. 1997, Reece et al 1997b, Saldarriaga et al. 2003, Villalba et al. 2004). This hypothesis was supported by serological affinities of P. marinus with some dinoflagellates including a few free-living as well as parasitic species of Hematodinium from Nephrops norvegicus, Chionoecetes bairdi, Portunus pelagicus, Callinectes sapidus, Necora puber and an unidentified gammaridean amphipod (Bushek et al. 2002a). Norn et al. (1999) proposed that perkinsids, which share features with both dinoflagellates and apicomplexans, be described as a taxon on level with other alveolate phyla, with the phylum name of Perkinsozoa (Robledo et al. 2011). Based on additional molecular analysis, Zhang et al. (2011a) supported the affiliation of the genus Perkinsus with an independent lineage (Perkinsozoa) positioned between the phyla of Apicomplexa and Dinoflagellata.

Geographic distribution

East coast of the United States of America (USA) from Maine to Florida (Andrews 1988,1996; Burreson and Ragone Calvo 1996; Ford and Tripp 1996), and along the Gulf of Mexico coast to the Yucutan Peninsula (Burreson et al. 1994a, Aguirre-Macedo et al. 2007, Gullian-Klanian et al. 2008). The prevalence and intensity of infection varies with location and season (Craig et al. 1989, Crosby and Roberts 1990, Oliver et al. 1998b). "Hot spots" of infection have been reported within this range (Brousseau et al. 1998) and Perkinsus marinus poses a threat to oyster aquaculture in some areas (Ulrich et al. 2007). The Maryland, USA, Department of Natural Resources, Fisheries Service monitors the prevalence, intensity and distribution of P. marinus in Chesapeake Bay (Tarnowiski 2003, 2005). The range extension of this disease into Delaware Bay, New Jersey and Cape Cod, Maine, USA in about 1990, was attributed to repeated introductions by many means over many years in conjunction with an increases in sea-surface temperatures particularly during the winter (Ford 1996, Cook et al. 1998, Ford et al. 2000a&b, Karolus et al. 2000, Ford and Chintala 2006, Powell et al. 2008, Pecher et al. 2008). In the more northern locations, temperatures for parasite proliferation (higher than 20C) are usually only suitable from June through September. Colder winters and high rainfall after 2002 reduced the prevalence of infection in some regions, but P. marinus can survive low temperatures and low salinities, and epizootic conditions are likely to return if temperatures rise again, as predicted by climate-change models (Ford and Smolowitz 2007). Perkinsus marinus was accidentally introduced into Pearl Harbor, Hawaii (Kern et al. 1973). In 2007, P. marinus was detected in cultured populations of endemic oysters (Crassostrea corteziensis) from the state of Nayarit on the Pacific coast of Mexico and these infections were associated with uncontrolled introductions of C. virginica in the region (Cceres-Martnez et al. 2008, 2010). Enrquez-Espinoza et al. (2010) attributed massive mortalities of farmed Crassostrea gigas in the Gulf of California (northwest Mexico) in July and August 2006 to P. marinus along with environmental factors. A Perkinsus-like parasite was reported in Crassostrea angulata imported into Great Britain from Portugal in the spring of 1969 (Alderman and Gras 1969). Perkinsus marinus as well as other species of Perkinsus were reported from Crassostrea rhizophorae, Crassostrea gasar and Crassostrea brasiliana in Brazil (Da Silva et al. 2012)

Host species

Crassostrea virginica, Crassostrea corteziensis and probably Crassostrea rhizophorae, Crassostrea gasar and Crassostrea brasiliana. Crassostrea gigas and Crassostrea ariakensis were also infected by experimental and environmental exposure but these two species seem to be more resistant to the disease (Barber 1996, Chu 1996, Calvo et al. 1999, Calvo et al. 2001, Paynter et al. 2008). Schott et al. (2008) reported that during experimental field trials, the prevalence of P. marinus was equivalent to or higher in C. ariakensis than in native C. virginica and in the laboratory, C. ariakensis efficiently transmitted the parasite to uninfected C. virginica. Crassostrea rhizophorae was also experimentally infected and seemed to be as susceptible to infection as C. virginica but may be somewhat more tolerant to heavier parasite infections (Bushek et al. 2002c). Cceres-Martnez et al. (2008) reported a relatively low prevalence of natural infections (less than 6%) with no associated mortalities in two farmed populations of Crassostrea corteziensis from the Pacific coast of Mexico. Clams, Mya arenaria and Macoma balthica were also experimentally susceptible to infection by mantle cavity inoculations (Dungan et al. 2007). Coss et al. (2001) detected P. marinus DNA in the clam Macoma mitchelli. However, surveys, using sensitive and specific molecular assays, for P. marinus in clams (n=452 of five species) sympatric (cohabitation) to C. virginica populations with high prevalences of P. marinus (50 to 100% infected) revealed P. marinus in only one clam (Mya arenaria) (Reece et al. 2008). Pecher et al. (2008) also reported that the prevalence of P. marinus in Mercenaria mercenaria was significantly lower than in C. virginica suggesting that M. mercenaria is not an optimal host for P. marinus. Aquacultured hard clams, M. mercenaria from the Gulf of Mexico coast of Florida, USA were negative for P. marinus by specific DNA probes (PCR) but low level infections were detected by Ray's thyoglycollate test and by genus-specific PCR (McCoy et al. 2007). Perkinsus marinus was also reported from the ectoparasitic snail Boonea impressa which was reported to transmit P. marinus between C. virginica in the laboratory (White et al. 1987). Note that C. virginica can also be infected by other species of Perkinsus (Coss et al. 2001, Pecher et al. 2008). In addition, two other species of Perkinsus (P. mediterraneus and P. beihaiensis) have been described from other species of oysters from European and Asian waters, respectively. Also, similar Perkinsus spp. have been reported from the rock oyster Saccostrea forskali (Taveekijakarn et al. 2005) as well as at least 34 species of molluscs (including clams, abalone and scallops) from warm temperate to tropical waters of the Atlantic and western Pacific oceans and Mediterranean Sea, from 30 species of bivalves on the Great Barrier Reef, and from cultured Pinctada maxima (pearl oysters) in Torres Strait, Australia (Norton et al. 1993, Perkins 1996). Mussels such as Mytilus edulis and Geukensia demissa, not known to develop perkinsosis, contain high anti-P. marinus activity in their haemolymph (Anderson and Beaven 2001).

Impact on the host

The endoparasite Perkinsus marinus has caused mass mortalities of C. virginica and is one of the primary factors that adversely impacts the abundance and productivity of this oyster at various locations along the east and Gulf coasts of the USA (Burreson and Ragone Calvo 1996, Ford 1996, Ray 1996, Powell et al. 2008). For example, the large decrease in oyster production in the Maryland part of Chesapeake Bay between 1981 and 1988, with landings as low as 15,000 metric tons since 1986 (compared to landings that fluctuating around 80,000 metric tons between about 1910 and 1980) was attributed to high mortalities related to P. marinus and Haplosporidium nelsoni, predation and poor management practices (Hral et al. 1990, Goulletque et al. 1994). However, the significant declines in C. virginica production in this area prior to 1950 was probably attributed to the mechanical destruction of habitat and stock over-fishing and not disease (Rothschild et al. 1994). Carnegie and Burreson (2008) noted that in the Virginia part of Chesapeake Bay, C. virginica populations persisted despite being infected with P. marinus and have a capacity for growth if substrate is managed effectively and the loss of reefs arrested. In Delaware Bay, C. virginica populations that are highly susceptible to P. marinus survive in the upper bay refugia possibly because environmental conditions inhibit infection (Pydeski and Bushek 2008). Gullian-Klanian et al. (2008) indicated that in Mexico, no serious epizootics attributed to P. marinus have been reported. Although high prevalences (mean of 70%) occurred in Terminos Lagoon (Mexico, Gulf of Mexico) during the dry season, the intensity of infection was usually light. Klanian et al. (2008) suggested that freshwater input associated with high nutrient concentrations during the rainy and north-wind seasons had a strong negative effect on P. marinus prevalence (23% and 7%, respectively) and also influenced the physiology of the oysters and that this seasonal stress was responsible for the absence of an epizootic event in Terminos Lagoon.

The life cycle of P. marinus includes trophozoites in the water column entering the paleal cavity of the oyster during filter-feeding and subsequently being directed through gills and palps towards the mouth. Once in the paleal cavity or the digestive tract, trophozoites displaying surface ligands for the oyster galectin CvGal (Tasumi and Vasta, 2007 get paper) are recognised and phagocytosed by the haemocytes that can transmigrate to the internal milieu and eventually into the vascular system. Parasites remain inside phagosome-like vesicles where they remain viable and multiply (tomonts). When infected haemocytes disintegrate, the released trophozoites can either be phagocytosed by neighbouring haemocytes or multiply extracellularly (tomonts) in both the internal milieu and lumen of the vascular system. The infected circulating haemocytes migrate throughout the host tissues where they lyse and release trophozoites, leading to systemic infection and eventually host death. Trophozoites are released into the water from live oysters via the feces and/or the pseudofeces and upon death of the oyster, from the decaying infected tissues (Bushek et al., 2002b). Once released into the water column, trophozoites may sporulate: trophozoites enlarge (prezoosporangia), develop a discharge tube and after multiple rounds of division, release hundreds of zoospores into the water column. Whether zoospores develop into trophozoites remains an open question (Robledo et al. 2011).

The effects of P. marinus infection in C. virginica range from pale appearance of the digestive gland, reductions in condition index, impaired gametogenic development, reduced haemolymph protein concentrations and lysozyme activity, to severe emaciation, gaping, shrinkage of the mantle away from the outer edge of the shell, retarded growth and occasionally the presence of pus-like pockets (Ford and Tripp 1996). The impact of P. marinus at varying infection intensities on the energy budget of the oyster can be estimated and explained with the acquisition of energy by P. marinus production and respiration (Choi et al. 1989). Remacha-Trivio et al. (2008) determined that the mean total number of trophozoites in eight naturally infected C. virginica was about 11.5 million cells with about 97% as mature trophozoites (signet rings), 2% as immature trophozoites (meronts) and 1% as clusters of small immature trophozoites (merozoites). Ford et al. (1999) reported that P. marinus distributions within host populations were aggregated (for example, 1 or 2 oysters may contain more parasites than all other oysters in the sample). Trophozoites occurred in the following tissues: intestine (30.1%), vesicular connective tissue (21.3%), haemocytes (14.9%), digestive gland (11.4%), gills (6.1%), connective tissues (5.7%), gonads (4.1%), palps (2.2%), muscle (1.9%), mantle connective tissue (0.8%), pericardium (0.7%), mantle epithelium (0.1%), and heart (0.1%) (Remacha-Trivio et al. 2008). Proliferation of the parasite causes systemic disruption of connective tissue and epithelial cells and is correlated with warm summer water temperatures (higher than 20 C) when pathogenicity and associated mortalities are highest. In South Carolina, the prevalence and intensity P. marinus in C. virginica was highest during periods of oyster spawning which also corresponded with warmer temperatures (Burrell et al. 1984, Bobo et al. 1989). However, in this area, relationships between salinity, temperature and gonadal stage of development of the oyster to P. marinus were not evident (Burrell et al. 1984). Nevertheless, differences in salinity were found to effect the prevalence and intensity of P. marinus in C. virginica in Chesapeake Bay (Paynter et al. 1993a). Also, in the mid northern region of the Gulf of Mexico, the intensity and prevalence of infection was correlated with salinity (Mackin 1955, Soniat et al. 2012) but not with water temperature (Soniat and Gauthier 1989, Gauthier et al. 1990). Infected C. virginica can eliminate viable P. marinus with the faeces and pseudofaeces at a rate correlated to both P. marinus body burden and subsequent survival time. However, in an epizootic, shedding of P. marinus via faeces is relatively small compared to the potential number released by cadavers of heavily infected oysters but may be important in transmission before infections become lethal (Bushek et al. 2002b). Marine aggregates (small clumps of material suspended in the water column) that contained P. marinus produced higher infection intensities than freely suspended P. marinus (Ralph et al. 2008) and important tissue sites of infection were the pallial organ (labial palps) and the principal pseudofaeces discharge area of the mantle (Winnicki et al. 2008). Hoese (1964) reported that snails, crabs and fish that scavenge on dead oyster tissues may serve as vectors of P. marimus after passive transfer of the parasite through the gut of the scavenger.

Saunders et al. (1993) determined that the growth rate of P. marinus in C. virginica was dependent on P. marinus population density and that this parasite could control its own population level but the delicate balance could be destabilized resulting in epizootics. Some C. virginica may survive summer proliferation of the parasite but are unable to revive following over-wintering dormancy. Mortalities of up to 95% have occurred in C. virginica during the second summer following transfer to disease enzootic areas. Although some researchers have reported inhibition of gonadal development and decline in reproductive output in infected oysters, Dittman et al. (2001) found that in Delaware Bay, the very drastic negative effect on reproduction predicted during the gametogenic phase did not materialize at the time of spawning because a period of diminishing parasite burdens coincided with gamete maturation. This observation was confirmed for C. virginica in Chesapeake Bay by Carnegie and Burreson (2011a). However in laboratory experiments, Chintala et al. (2002) found increased levels of infection in oysters undergoing spawning and/or exposed to low oxygen stress. Willson and Burnett (2000) did not find a strong correlation between oxygen uptake and infection intensity with P. marinus at either 25 or 35 C. However, Breitburg et al. (2011) reported that diel-cycling hypoxia increased acquisition of P. marinus infections, most likely by reducing the oyster defence responses but decreased the release of infective stages possibly because of lower filtration rates and consequently lower faeces production.

Perkinsus marinus incorporates and modifies lipids and synthesises phospholipids from exogenous sources and the metabolic modes of trophozoites (meronts) differ from those of prezoosporangia (Chu et al. 1998, 1999; Lund and Chu 2002). Unlike other protistan parasites, trophozoites of P. marinus are capable of synthesizing long chain essential fatty acids de novo (specifically, unsaturated fatty arachidonic acid 20:4(n-6) from acetate) in vitro via a delta-8 pathway (Chu et al. 2002, 2004). Venegas-Calern et al. (2007) examined the genetics associated with this process. Perkinsus marinus is an agent of stress on its host as assessed by the biochemical measurement of the taurine-to-glycine ratio in C. virginica (Soniat and Koenig 1982). Infection decreased taurine and glycine levels by about 40% and decreased free amino acid levels by about 33% thereby possibly impairing the salinity tolerance mechanisms of C. virginica (Paynter et al. 1993b, 1995). Although haemocytes from infected C. virginica had a greater zymosan-induced chemiluminescence (Anderson et al. 1992, 1993, 1995), P. marinus trophozoites did not stimulate a chemiluminescence response in oyster haemocytes (Anderson 1999a). This suggests that P. marinus may be able to suppress or remove the release of reactive oxygen intermediates from haemocytes, thus evading this component of the host's defence despite the capability of the oyster haemocytes to recognize and phagocytize P. marinus (Volety and Chu 1995, La Peyre et al. 1995b, Anderson 1999b, Schott et al. 2003a). One mechanism that may be employed by P. marinus to abrogate the respiratory burst by haemocytes (the host's oxidative defence response) following phagocytosis is the dismutation of superoxide radicals to molecular oxygen and hydrogen peroxide by superoxide dismutases (SODs) identified by Asojo et al. (2006) in P. marinus as iron-cofactored PmSOD1 and PmSOD2. In vitro challenges showed that haemocytes isolated from C. virginica can kill about 50% of P. marinus from cultured isolates (Volety and Fisher 2000). However, P. marinus produces many extracellular proteins (ECP) in vitro and there is evidence that these ECP, especially proteases, are important in vivo for the establishment of infection, defence parameters of the oyster and propagation of the parasite (Garreis et al. 1996; La Peyre and Faisal 1997a; Volety and Chu 1997; La Peyre and Cooper 1998; Oliver et al. 1998a, 1999b, b; La Peyre and Volety 1999; Ottinger et al. 2001; Wright et al. 2002; Ahmed et al. 2003; Muoz et al. 2003; Schott and Vasta 2003;  Schott et al. 2003b; Jordan and Deaton 2005). Also, EPC activity is affected by temperature (Chu et al. 2003) and suppresses the vibriocidal activity of oyster haemocytes to effectively eliminate the bacterium Vibrio vulnificus, potentially leading to conditions favouring higher numbers of vibrios in oyster tissues (Tall et al. 1999). Robledo et al. (2004) identified the complementary DNA (cDNA) sequence of the divalent cation transporter natural resistance–associated macrophage protein homologue from P. marinus (PmNramp) that may relate to the parasite’s survival of intracellular killing by the host.

Antimicrobial activity against P. marinus occurs in cell-free haemolymph (plasma) of oysters (Anderson and Beaven 2000). Although serum agglutinins were not associated with resistance to P. marinus (Chintala et al. 1994), low molecular weight protein inhibitors (such as the slow-tight binding serine protease inhibitor described by Xue et al. (2006)) and high antiproteolytic activity in the haemolymph of C. virginica and C. gigas may play a role in oyster host defence mechanisms (Oliver et al. 1999a, 2000; Faisal 1999; Elsayed and Faisal 1999; Xue et al. 2006). Chu and La Peyre (1989) found no linkage between hemolymph lysozyme and protein concentration and infection of oysters by P. marinus. However, C. virginica at low temperature and low salinity had higher plasma lysozyme concentrations which may result in an unfavourable environment for the development of the parasite and/or weaken parasite activity (Chu and Volety 1997a, Chu 1998). La Peyre et al. (2010) indicated that in vitro, the largest decreases in P. marinus viability occurred only with combined low temperature and low salinity, indicating that there is clearly a synergistic effect between these environmental parameters on the parasite. The role (if any) of heat shock proteins in stress and disease resistance in oysters requires further investigation (Brown et al. 1993, Tirard et al. 1995, Encomio and Chu 2005). However, the heat shock response in C. virginica was not negatively affected by P. marinus infection (Encomio and Chu 2007). Wang et al. (2010) employed microarray analysis of gene expression in C. virginica to reveal a novel combination of antimicrobial and oxidative stress host responses 30 days after challenge with P. marinus.

Bushek and Allen (1996a) found that races of C. virginica resistant to P. marinus exist and that natural levels of resistance roughly corresponded to the extent ancestral populations had been exposed to P. marinus. Also, differential selection by P. marinus can generate divergence in popluation structure in C. virginica (Green-Beach et al. 2011). Sokolova et al (2006) found two arbitrary fragment length polymorphism (AFLP) markers in C. virginica from North Corolina, USA, that were associated with infection and suggested the existence of genes or groups of genes that act directly or indirectly to control the levels of infection. Bushek and Allen (1996b) also indicated that races of P. marinus vary in virulence and environmental tolerance. The occurrence of different P. marinus strains with distinct phenotypic traits was confirmed by Yee et al. (2005) and microsatellite analysis of P. marinus genotypes from Florida and New Jersey indicated limited parasite migration between geographically proximate populations of oysters (Thompson et al. 2008). Reece et al. (2001a, b) detected 12 different composite genotypes of P. marinus with >88% of 76 isolates possessing one of three predominant genotypes but they found that a single oyster can be infected with multiple strains.

Modeling of diseased C. virginica populations by Hofmann et al. (1995) and Powell et al. (1996) showed that food availability to the host and salinity and temperature control on the growth and development rates of the host and parasite are environmental factors involved in regulating P. marinus. Temperature, salinity and their interaction are important environmental influences on transmission and pathogenicity of P. marinus in C. virginica (Crosby and Roberts 1990, Audemard et al. 2006). However, other environmental and biological factors (e.g., pollution, other human influences, host nutrition and growth, spawning and reproduction, age, resistance, oyster density and distribution and disease vectors) affect the levels of parasitism observed in the field (Craig et al. 1989, Soniat 1996). For example, Ragone Calvo et al. (2003b) suggested that atypical early-summer oyster mortality from Haplosporidium nelsoni, at a time when infections of P. marinus are light, can have a significant indirect influence on P. marinus transmission dynamics. Also, a relationship with changes in salinity caused by climatic cycles was associated with changes in disease prevalence and intensity (Soniat et al. 2006, 2009, 2012). Lenihan et al. (1999) and Volety et al. (2000) demonstrated that the depth below mean low water at which the oyster occurs on the reef affects the prevalence and intensity of infection and oyster mortality with the most significant impact on oysters towards the base of the reef. Also, land use patterns (anthropogenic disturbances) may affect the distribution of the disease and exacerbation of oyster mortalities (White et al. 1998, Bushek et al. 2000b, Power et al. 2006). For example, in the USA, Gulf of Mexico, Wilson et al. (1990, 1992) reported that latitude, total petrolium aromatic hydrocarbon (PAH) content and industrial and agricultural land use significantly affected the parasite's distribution and concluded that industrial and agricultural activity may correspond with high prevalence and intensity of disease caused by P. marinus.

Exposure of the oyster to pollutants such as the chemical carcinogen n-nitrosodiethylamine (DENA) and tributyltin (TBT) will enhance the disease caused by P. marinus (Winstead and Couch 1988 and Anderson et al. 1996, respectively). Also, exposure of C. virginica to field contaminated sediments and related water soluble fractions for about one month in the laboratory significantly elevated the expression/ progression of latent P. marinus infections in a dose-dependent manner (Chu and Hale 1994, Chu 1999). However, Burreson and Ragone Calvo (1996) claim that there is little evidence to support the common perception that pollution is responsible for the dramatic increase in P. marinus abundance since 1985. An assessment of the impact of several common anthropogenic contaminants on the proliferation of P. marinus in vitro determined that only a herbicide (with active ingredients of 3.1% 2,4-dichlorophenoxyacetic acid, 10.6% mecoprop and 1.3% dicamba that mimic growth hormones of broadleaf plants to over stimulate growth resulting in plant death) had a significant negative effect, but only above the manufacture’s recommended application rate (Bushek et al. 2007).

Crassostrea gigas and triploid C. ariakensis were found to be less susceptible (but not completely resistant) to infection and disease caused by P. marinus (Meyers et al. 1991; Calvo et al. 1999, 2000, 2001 ). However, C. gigas may be less tolerant to environmental factors prevailing in usual C. virginica habitat (Barber and Mann 1994, Chu 1996). Greater resistance to infection and disease in C. gigas may be attributed to elevated cellular and humoral activities (including protease inhibitors) that may degrade the parasite more effectively, and/or lower plasma protein levels that may limit parasite growth (La Peyre et al. 1995a, Romestand et al. 2002, Goedken et al. 2005). Also, differences in gene expression in challenged C. virginica and C. gigas were identified by suppression subtractive hybridization (Tanguy et al. 2004). Although sterile triploid C. virginica grew faster and had a lower cumulative mortality than diploids, the triploids were equally susceptible to P. marinus (Barber and Mann 1991, Dgremont et al. 2012). Advanced infections observed in C. ariakensis exposed to P. marinus in the laboratory suggests that there may be some risk of mortality to C. ariakensis if this oyster is held under stressful conditions at least in hatchery or laboratory settings (Moss et al. 2006)

Diagnostic techniques

Gross Observations: Infected oysters may have a pale appearance to the digestive gland, reductions in condition index, severe emaciation, gaping, shrinkage of the mantle away from the outer edge of the shell, reduced gonadal development and/or retardation of growth. Occasionally, pus-like pockets may occur in the soft tissues. However, these signs are not pathognomonic of perkinsosis. Cceres-Martnez et al. (2008) reported that infection in C. corteziensis was associated with gross signs of weakness and transparent and retracted mantle.

Wet Mount: Spherical bodies containing an eccentric vacuole ("signet-ring") in preparations from moribund oysters. Because other species of Perkinsus share this morphological feature, other methods will have to be employed to determine the specific identity of organisms. Cheng and Manzi (1996) indicated that P. marinus could be detected in haemolymph of infected oysters following the application of a "panning" technique (specifically, about 1 milliliter of haemolymph was placed on the bottom of a Petri dish, incubated at 25 C for 30 minutes followed by examination of the nonadhering cells for parasites).

Histology: Because the infection is usually systemic, the connective tissue of all organs may harbour immature trophozoites (= meronts, merozoites or aplanospores, 2-3 m in diameter), mature trophozoites (= "signet-ring" stages or mature meronts, merozoites, trophonts or aplanospores, 3-10 m in diameter each containing a large eccentric vacuole (possibly containing a vacuoplast) that presses the nucleus to the periphery of the cell) and tomonts (="rosettes", sporangia or schizonts, 4 -15 m in diameter and containing 2, 4, 8, 16 or 32 developing immature trophozoites). Many of the parasites may occur within oyster haemocytes. Trophozoite morphology does not have taxonomic value because it can be influenced by the host, the time of the year, and nutrient availability (Villalba et al. 2004). For example, Carnegie and Burreson (2011b) reported that since 1986, large trophozoites of P. marinus at relatively low infection intensities in C. virginica from Chesapeake Bay have been replaced by abundant minute parasite cells. Histopathology of severe infections in C. corteziensis, as well as other susceptible oysters, consists of general invasive infiltration of haemocytes, including phagocytosis of parasite stages, that were disseminated in the connective tissue surrounding the epithelia of the digestive gland, gonad and mantle with the presence of some brown cells (Cceres-Martnez et al. 2008).

Figures 1 to 3. Perkinsus marinus in histological sections of the connective tissue in the digestive gland (Figs. 1 to 3) and intestinal epithelium (Fig. 4) of Crassostrea virginica from Maryland, USA. Haematoxylin and eosin stain.

Histological sections

Figure 1. Two mature trophozoites ("signet ring" stage). The large eccentric vacuole (V) and nucleus (N) are indicated on the specimen closest to the wall of a digestive gland tubule.

Histological sections

Figure 2. Mature trophozoites (M) in which the basophilic chromatin of the nucleus appears as a ring around the perimeter of the nucleus, and two eight-cell tomonts (T) within which immature trophozoites are developing.

Histological sections

Figure 3. A 16-cell tomont (T) containing developing trophozoites. This tomont is contained within a haemocyte (HN indicates the nucleus of the phagocytic cell) and a maturing trophozoite (M) is located near by.

Histological sections

Figure 4. Heavy infection consisting of trophozoites (Z), tomonts (T) and mature trophozoites (M).

Electron Microscopy: A marked characteristic of all species of Perkinsus is the ultrastructure of the zoospore. For P. marinus, zoospores have been produced by transferring the prezoosporangium from thioglycollate medium (see cultures below) into sea water where it develops into a zoosporangium that produces hundreds of motile biflagellate zoospores. The anterior flagellum is ornamented with hair-like structures (mastigonemes) and spurs-like structures and the posterior flagellum is glabrous. The zoospore contains an apical complex consisting of a conoid, subpellicular microtubules, rhoptries, rectilinear micronemes and conoid-associated micronemes. Large vacuoles also occur at the anterior end of the zoospore. (For ultrastructural details and illustrations of zoospores see Perkins 1976b and Perkins 1996). Perkins (1969) described the ultrastructure of vegetative stages as found in the host C. virginica. Sunila et al. (2001) described the ultrastructural characteristics of P. marinus vegetative stages (trophozoites (=trophonts) and tomonts (=schizonts)) from in vitro propagation cultures, and zoosporangia isolated from C. virginica haemolymph, enlarged in Fluid Thioglycollate Medium (as described below) and zoosporulated in Dulbecco's Modified Eagle/Ham's F-12 liquid medium supplemented 3% fetal bovine serum and antibiotics.

Immunological Assay: Polyclonal antibodies that recognized only prezoosporangia (=hypnospores) of P. marinus (Choi et al. 1991) and others that bind to most life stages of many but not all species of Perkinsus as well as other parasitic dinoflagellates and monoclonal antibodies that recognize epitopes unique to the prezoosporangia have been produced (Dungan and Roberson 1993, Bushek et al. 2002a). Other monoclonal antibodies produced by Romestand et al. (2001) detected P. marinus trophozoites and their protein lysates and also trophozoites from Perkinsus olseni (=atlanticus) from European clams (Ruditapes decussatus). Immunoassays have been successfully applied to various environmental samples and in vitro experimental systems (Dungan 1997). Polyclonal antibodies produced to purified P. marinus extracellular proteins recovered from defined culture medium were used to produce an ELISA-based assay that may be comparable to Ray's thioglycollate test as described below (Kaattari et al. 1997, Dungan and Hamilton 1997, Ottinger et al. 2001). Remacha-Trivio et al. (2008) used stereological methods (defined as a collection of strongly based mathematical procedures aimed to quantify geometrical properties of target objects without assumptions concerning inherent characteristics of these objects) and immunohistochemistry (using the rabbit anti- P. marinus IgG of Dungan and Roberson (1993)) to quantify and determine the tissue distribution of P. marinus in C. virginica.

DNA Probes: Small subunit ribosomal RNA gene (SSU rDNA, at least 1,793 nucleotides) has been sequenced by polymerase chain reaction (PCR) and molecular cloning. Specific and sensitive semiquantitative and competitive PCR assays, including multiplex PCR (simultaneous testing of two or more pathogens in a single test reaction), and quantitative real-time PCR assays were developed for: 1) assessing intensity (including quantification) of P. marinus in oysters, 2) determining prevalence of infections in oyster populations, 3) monitoring of over-wintering oyster populations, 4) certifying disease free oyster spat, 5) elucidating mechanisms of infection, 6) assessing the presence of the parasite in environmental waters and various invertebrate species and 7) differentiating between various species of Perkinsus (Marsh et al. 1995; Vasta et al. 1997; Penna et al. 1999, 2001; Russell et al. 2000, 2004; Elandalloussi et al. 2004; Abollo et al. 2006; Audemard et al. 2004, 2006; Pecher et al. 2008). Robledo et al. (1998), Yarnall et al. (2000) and Gauthier et al. (2006) described various PCR based assays to detect P. marinus that proved to be more sensitive than Ray's thioglycollate test (described below). Inter- and intra-specific genetic variation among Perkinsus species has provided the opportunity to design genus- and species-specific molecular diagnostic assays (Reece et al. 2001a, b). However, before molecular analysis (e.g., PCR) can be recommended as the method of choice for disease diagnosis, more research is necessary to validate the various molecular diagnostic assays and compare them to standard diagnostic techniques (Burreson 2000, Reece and Burreson 2004, Villalba et al. 2004). Apparently, the real-time quantitative PCR assay developed and validated by De Faveri et al. (2009) correlated with traditional detection methods (Ray's thioglycollate test) and did not amplify Perkinsus chesapeaki or Perkinsus olseni DNA.

Molecular sequence data is playing an increasingly important role in the identification of Perkinsus species and requires adequate DNA sequence data at the targeted loci from the same and related species over a wide geographic area in order to develop reliable, accurate and sensitive molecular diagnostic tools (Villalba et al 2004). Thirteen percent difference in the internal transcribed spacer 1 (ITS1) plus internal transcribed spacer 2 (ITS2) of the SSU rDNA between P. marinus and 4 other isolates of Perkinsus (P. olseni (=atlanticus) from clams in Portugal, P. olseni from abalone in Australia, isolates of Perkinsus sp. from the jewel box and blood cockle in Australia) is a strong indication that P. marinus of C. virginica is a distinct species. Nevertheless, Brown et al. (2004) found 14 polymorphic nucleotide positions at the ITS region (8 in ITS1 and 6 in ITS2) in 12 isolates of P. marinus from the Atlantic and Gulf of Mexico coasts of the USA and indicated the importance of determining the genetic variation of each locus prior to development of sequence-based molecular diagnostics. The use of PCR primers to amplify up to six polymorphic loci of genomic DNA from cultured P. marinus indicated that in vitro P. marinus are diploid and that oysters may be infected by multiple strains of this parasite (Reece et al. 1997a, c, 1999). The serine protease gene in P. marinus is variable and may correlate with its virulence or pathogenicity (Brown and Reece 2000, 2001, 2003). Allele sequences were identified in isolates from geographically distant sites. However, allelic and genotypic frequencies differed significantly among isolates from regions of the northeast and southeast US Atlantic coast and the coast of the Gulf of Mexico (Reece et al. 1999). Primers that target the non-transcribed spacer (NTS), a region with high inter-specific variation, have demonstrated good species specificity (Robledo et al. 1998), although intra-specific variations were detected and the prevalence of the two described types varied with the geographic origin of the samples (Robledo et al. 1999). However, intra-specific variations within the NTS region has not been broadly assessed creating a risk of false negatives due to polymorphism within a species if the PCR primers do not bind the target sequence of all strains of that species (Villalba et al. 2004). Assessing high resolution microsatellite markers and amplified alleles directly from infected oyster genomic DNA, Thompson et al. (2011) suggested that P. marinus employs multiple reproductive modes, and that over the short term, selection acts upon independent parasite lineages rather than upon individual loci in a cohesive, interbreeding population. Nevertheless, high genotypic diversity is the evolutionary legacy of sex in P. marinus. Anthropogenic movement of infected oysters may increase outcrossing opportunities, potentially facilitating rapid evolution of this parasite (Thompson et al. 2011). Other components of the P. marinus genome have been described but to date, none of these have been developed into diagnostic assays (Zhang et al. 2011 a, b, Hearne and Pitula 2011, Robledo et al. 2011)

Culture: Examine tissues for blue-black prezoosporangia (= hypnospores, usually 30-80 m in diameter but extremes of 480 m in diameter have been observed), after incubation in Fluid Thioglycollate Medium (FTM) supplemented as described by Ray (1966) for approximately 7 days followed by staining with Lugol's iodine stain, (see Ray (1966), Choi et al. (1989), Fisher and Oliver (1996) and Kim et al. (2006) for details of the technique and see Nickens et al. (2002) for an alternative formulation). This diagnostic procedure is frequently referred to as Ray's thioglycollate test (or technique, Ray's FTM, RFTM). Although not true propagating cultures, this procedure is used for the diagnosis of many species of Perkinsus but may also detect other organisms (Villalba et al. 2004). The oyster tissue that is assayed for parasite surveys or monitoring work tend to reflect the same seasonal trend but the choice of tissue is important depending on the accuracy required if oyster body burden of the parasite is being assessed (Bushek et al. 1994, Oliver et al. 1998b). Methods to improve determination of the intensity of infection, (whole-oyster parasite burden) described by Fisher and Oliver (1996), have been suggested by Coates et al. (1999). Ray's thioglycollate test has been used to quantify the intensity of infection without sacrificing the oyster by determining the number of parasites in a haemolymph sample (Gauthier and Fisher 1990; Nickens et al. 2000b, 2002). The use of Ray's thioglycollate test to detect and quantify planktonic P. marinus in environmental water samples has been proposed by Ellin and Bushek (1999, 2000b, 2006). However, other methods will have to be employed to determine the specific identity of organisms that may stain with Lugol's iodine following incubation in FTM. Audemard et al. (2008) indicated that P. marinus DNA could be successfully amplified from samples processed by Ray's thioglycollate test including staining with Lugol's iodine stain. Thus, the identity of the Perkinsus sp. in a positive Ray's thioglycollate test can be determined with the added advantage of being able to preserve positive samples in 95% ethanol for later processing for DNA amplification and sequencing or submission to a laboratory with molecular identification capabilities (Audemard et al. 2008).

Wet Mount

Figure 5. Wet mount of the rectum of Crassostrea virginica that was processed by the Ray's thioglycollate test (including staining with Lugol's iodine) to reveal six enlarged and darkly (blue-black) staining prezoosporangium of Perkinsus marinus.

Methods for the in vitro propagation of histozoic stages of P. marinus were described by Kleinschuster and Swink (1993), La Peyre et al. (1993), Gauthier and Vasta (1993, 1995), La Peyre and Faisal (1995a, 1995b, 1996), La Peyre (1996), and Soudant and Chu 2001). La Peyre and Chu (1994) described a simple procedure for the isolation of P. marinus trophozoites (merozoites) from heavily infected C. virginica. The in vitro culture of this parasite has led to insights into its biology. Krantz (1994) used in vitro cultures to assay chemicals for inhibitory activity and to screen for potential chemotherapeutic agents. Perkinsus marinus cultured in nutrient media were sensitive to low salinity with mortalities increasing as salinity decreased below 12 ppt. By culturing P. marinus in media of various osmolarities (168 to 737 mOsm - equivalent to 6.5 to 27.0 ppt), O'Farrell et al. (2000) determined that the size of P. marinus vegetative stages varied with osmolarity and cells cultured at low osmolarity can withstand hypoosmotic (56 mOsm - 2.5 ppt) and hyperosmotic (672 mOsm - 24.7 ppt) stress (41% mortality at 2.5 ppt) where as cells cultured at high osmolarity experienced 100% mortality when transferred to the hypoosmotic artificial seawater. La Peyre et al. (2006) reported that P. marinus exhibited reduced viability at 7 ppt, even after acclimation. Lund et al. (2004) found that in culture media, salinity treatments (about 14, 20 and 28 ppt) exhibited few treatment effects, but temperature significantly affected cell proliferation, fatty acid content and fatty acid synthesis rates (i.e., fatty acid synthesis rates increased approximately two-fold for every 10 C increase in temperature). Ford and Chintala (2006) used in vitro data to test the hypothesis that the northward expansion of P. marinus was associated with a low-temperature adapted strain of the parasite. They found no evidence of low-temperature adaptation by P. marinus based on the fact that net proliferation rates for isolates were similar at temperatures from 5 to 20 C. However, at temperatures of 25 to 35 C, the South Carolina isolates exhibited higher proliferation rates than the northern isolates suggesting possible high-temperature adaptation of parasite strains that are routinely exposed to higher temperatures. La Peyre et al. (2008b) determined that P. marinus had 49% viability after 30 days at 4 C, but limited metabolic activity and no proliferation which could partially explain decreasing parasite infection intensities in C. virginica during the colder months of the year. Using in vitro assays, Soudant and Chu (2001) determined that during immature trophozoite proliferation, P. marinus synthesizes certain fatty acids and lipid classes, but for development from immature trophozoites to prezoosporangium, the parasite may rely on its host for lipid resources. For example, Lund et al. (2007) suggested that P. marinus cannot synthesize sterols and must sequester them from its host. Also, extracts or haemolymph from C. ariakensis, C. gigas and other non-oyster mollusc species significantly reduced in vitro proliferation compared to extracts from C. virginica (Gauthier and Vasta 2002, Brown et al. 2005). Tissue extracts from C. virginica caused P. marinus to secreted elevated amounts of a set of low molecular weight serine proteases (LMP: 30–45 kDa) which were not upregulated by extracts from C. gigas and C. ariakensis (MacIntyre et al. 2003).

Five percent fetal bovine serum was required in one culture media formulation but higher concentrations dramatically reduce parasite proliferation in a dose-dependant manner because of transferrin (a natural iron chelator) within the serum which sequesters available iron resulting in P. marinus growth inhibition (Gauthier and Vasta 1994). An in vitro tetrazolium-based cell proliferation assay was described for monitoring the affect of conditions on P. marinus multiplication (Dungan and Hamilton 1995). La Peyre and Faisal (1997b) described a protein-free chemically defined culture medium and proposed that the medium was suitable for the study of P. marinus proteins, to produce antigens for antibody production and to screen chemotherapeutic agents. Bushek et al. (2000a) described a technique to clone P. marinus using micromanipulation and a "feeder layer".

Perkinsus marinus produced in vitro were infective to oysters by injection into the shell cavity or adductor muscle but not via feeding. Ford et al. (2002) reported that virulence was lost immediately in culture and repassing cultured P. marinus through oysters did not restore virulence. Also, P. marinus in log-phase were significantly more virulent than those from lag- or stationary-phase cultures (Ford et al. 2002, Chintala et al. 2002). Although, cultured P. marinus appeared to have a low pathogenicity (Bushek et al. 1997a, 2002b), supplementing the cultures with oyster plasma or tissue homogenates seemed to enhance infectivity (Earnhart et al. 2004) and led to pronounced changes in P. marinus cellular morphology comparable to those observed within naturally infected oysters (MacIntyre et al. 2003). Shaheen (1999) determined the P. marinus can survive in artificial seawater at 22 ppt salinity and 27 C for more that 6 weeks without an exogenous supply of nutrients. Cryopreserved isolates of P. marinus are available at the American Type Culture Collection (ATTC, Rockville, MD, USA).

Methods of control

Oysters from areas with records of the disease should not be imported into Canada. Perkinsus marinus is easily transmitted between oysters, thus, it is imperative to avoid moving infected oysters to an area containing uninfected oysters (Ford and Tripp 1996). To date, eradication has proven impossible. Management methods used to reduce the commercial impact of the disease on infected populations consist of reducing the density of oysters and harvesting or moving oysters to low salinity areas (lower than 9 ppt (Ragone and Burreson 1993)) before water temperatures increase to 15-20 C (however, infections may persist for years in low salinity areas (Burreson and Ragone Calvo 1996)). From the results of in vitro experiments on P. marinus cultured at different salinities, O'Farrell et al. (2000) suggested that transferring infected oysters to low salinity will result in strains of P. marinus acclimated to low salinity and thus able to withstand periodic events of extremely low salinity. Thus, caution is necessary when using low salinity areas to treat or control infection and/or disease (Paynter and Burreson 1991). However, repetitive and well-timed low salinity (freshet) events can prevent infection or at least maintain P. marinus at non-lethal intensities making the control of freshwater inflows a possible adaptive management approach (Mackin 1955, La Peyre et al. 2003, 2009). Specifically, low salinity events (less than 5 ppt) decreased Perkinsus marinus infection intensities, even as temperatures exceeded 20C (La Peyre et al. 2009). This relationship between P. marinus and salinity was used by authorities in coastal areas of Texas as a biological indicator for determining minimum freshwater inflows (Culbertson et al. 2011). Nevertheless, Fisher et al. (1992) and Chu and Volety (1997b) found that temperatures (between 10 C and 28 C) was more influential than salinity (3 to 39 ppt) in affecting susceptibility to P. marinus in oysters, the intensity of P. marinus infections and oyster mortalities. The relationship between changes in salinity associated with climate change and changes in disease prevalence and intensity suggested an approach for predicting epizootics of P. marinus from climate models and thus be used in the management of oyster populations (Soniat et al. 2006).

Chu and Greene (1989) used prezoosporangia and zoospores obtained from Ray's thyoglycollate test to determine that prezoosporangia cannot withstand temperatures as low as 4 C for more than 4 days and zoospores died in 1 day when transferred from 28 C to 4 C. Cold winter temperatures may limit the natural spread of this pathogen to northern areas (infection declines at temperatures below 15-20 C). However, oysters held for 11 weeks at 15 C, a temperature considered more favourable for oyster haemocytes than for P. marinus, were not able to eliminate infections (Ford et al. 1999). In enzootic areas, strategies designed to enhance and supplement natural recruitment of oysters, along with keeping growing areas free from P. marinus by limiting oyster transplantation, currently offer the most promise for maintaining commercially harvestable stocks (Krantz and Jordan 1996). Paynter et al. (2010) determined that hatchery-produced juvenile oysters planted on numerous natural oyster bar in Maryland between 1995 and 2009 grew well enough to reach market size in 2 to 3 years and the rates of P. marinus infection were low. Thus, they suggested that disease-related mortality would not often threaten oyster aquaculture within the area (Paynter et al. 2010). Management strategies of fallowing beds after removing infected oysters has not proven effective in some areas but early harvest to avoid mortalities caused by P. marinus may be feasible (Butsic et al. 2000). La Peyre et al. (2008a) suggested that oysters cultured off-bottom on adjustable long-line systems and exposed to air daily had increased survival and higher condition indices. Ford et al. (2000c, 2001) demonstrated that juvenile oysters (seed) from nursery systems that use raw water pumped from an enzootic area are highly likely to be infected although infections may be very light in intensity and low in prevalence. Treating raw water, by filtration to 1 m and then exposure to ultraviolet light (30,000 W s-1 cm-2 UV irradiation), will help protect hatchery produced seed from infection (Ford et al. 2001). Deploying specific-pathogen-free C. virginica into enzootic areas may not reduce infection and subsequent mortalities, and ultimate success with production will depend on the salinity regime they experience during grow-out (Albright et al. 2007).

Efforts to identify Crassostrea virginica stocks resistant to Perkinsus marinus are in progress (Bushek and Allen 1996a, b; Stickler et al. 2001; Encomio et al. 2005, Wang and Guo 2008). Some disease-tolerant strains of C. virginica had better growth and survival than other strains (Abbe et al. 2010) and wild oysters from disease prone locations could contribute to the development of disease resistance in farmed oysters (Roberts et al. 2008). Although, stocks selected for resistance to other pathogens (i.e., Haplosporidian nelsoni) were highly susceptible to P. marinus (Burreson 1991), dual resistance to both parasites was achieved through four generations of artificial selection at a location where both diseases are enzootic (lower York River, Virginia, USA) (Ragone Calvo et al. 2003a). Also, interline crossing of C. virginica stocks developed for resistance to P. marinus, H. nelsoni and Roseovarius sp. seemed to perform well under a variety of disease pressures (Rawson et al. 2008). Powell et al. (2011) used a gene-based population dynamics model to assess the apparent limited development of resistance by C. virginica to P. marinus and determined that a mortality rate that limits the development of disease resistance still strains the ability of the species to maintain a vibrant population necessary to its long-term survival. Specifically, a limited infusion of susceptible larvae might be sufficient to offset any selective advantage realized from an epizootic mortality rate of 20 to 25%.

Based on the results of in vitro and in vivo studies, Faisal et al. (1999) suggested that bacitracin (a family of branched cyclopeptide antibiotics produced by Bacillus licheniformis and B. subtilis and used to treat various bacterial and protozoan infections in vertebrates) has promise for use in P. marinus chemotherapy. Lund et al. (2005) and Chu et al. (2008) suggested that the antimicrobial drug triclosan (5-chloro-2-(2,4 dichlorophenoxy) phenol, a specific inhibitor of Fab1 (enoyl-acyl-carrier-protein reductase) an enzyme in the Type II class of fatty acid synthetases) may be effective in treating P. marinus-infected oysters. However, quinine (a traditional preventative and treatment for the malarial parasites Plasmodium spp.) were lethal to infected oysters at concentration below the effective concentrations for P. marinus trophozoies (meronts) in vitro or had no observable effect on parasite infections (Panko et al. 2008).

The standard bleach sterilization procedures which use chlorine concentrations of 300 parts per million for 30 minutes as a method to kill P. marinus, prior to ocean disposal of P. marinus contaminated materials, are ineffective according to Bushek et al. (1997b and c). The quarantine of oyster shells on land for at least one month can dramatically reduce the potential risk of spreading P. marinus when using oysters shells (e.g., as cultch) from contaminated areas in the marine environment at other geographic locations (Bushek et al. 2004). Apparently, one hour at 50 C or one hour exposure to fresh water effectively killed the parasite (Bushek et al. 1997b). Organic N-halamine disinfectants (up to 25 mg/L for up to 12 h exposure, depending on the specific chemical formulation) can also be used to disinfect seawater contaminated with P. marinus (Delaney et al. 2003).

Seasonal proliferation of P. marinus has been modeled to estimate time to critical levels and duration of infection in C. virginica (Hofmann et al. 1995, 1999; Soniat and Kortright 1998; Brewster et al. 1999, 2000; Ragone Calvo and Burreson 2000; Brousseau and Baglivo 2000a, b; Ragone Calvo et al. 2001). One model has been developed into an internet program that may assist in calculating the time to a critical level of disease (Soniat et al. 2000, Ray et al. 2001; see advertisement by Scarratt 2000). Simulations of the model developed by Hofmann et al. (1999) could be used to understand the causes underlying the northward spread of the disease and to restructure the practices of the oyster industry to maximize production under conditions where the life span of the commercial species is controlled by disease. The model of Ragone Calvo et al. (2001) suggests that a single transmission event may be sufficient for P. marinus to become enzootic in a specific year class of oysters located in moderate to high salinity areas, while periodic transmission events are required for the parasite to persist in low salinity areas. Field studies by McCollough et al. (2007) confirmed the development of epizootics within 8 weeks of exposure to local infection pressures when first infections were simultaneously detected among greater than 62% of the specific-pathogen-free juvenile C. virginica experimentally deployed into a mesohaline enzootic area. A model for managing the oyster fishery during times when disease is a controlling influence was developed and assessed using oyster populations affected by P. marinus (Klink et al. 2001). Jordan (1995) used cluster (multivariate classification) analysis to evaluate oyster population structure and disease dynamics of some Maryland C. virginica populations.

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Citation Information

Bower, S.M. (2011): Synopsis of Infectious Diseases and Parasites of Commercially Exploited Shellfish: Perkinsus marinus ("Dermo" Disease) of Oysters.

Date last revised:  February 2011
Comments to Susan Bower